Hostname: page-component-78c5997874-xbtfd Total loading time: 0 Render date: 2024-11-05T11:33:17.600Z Has data issue: false hasContentIssue false
  • English
  • Français

Molecular genetic basis of flower colour variegation in Linaria

Published online by Cambridge University Press:  26 September 2007

LISETE GALEGO  and
JORGE ALMEIDA
LISETE GALEGO
Affiliation:
Division of Plant Science, Instituto de Tecnologia Química e Biológica, Oeiras, Portugal
JORGE ALMEIDA*
Affiliation:
Division of Plant Science, Instituto de Tecnologia Química e Biológica, Oeiras, Portugal Departamento de Botânica e Engenharia Biológica, Instituto Superior de Agronomia, Lisbon, Portugal
*
*Corresponding author. Instituto de Tecnologia Química e Biológica, Av. da Republica, 2781-901, Oeiras, Portugal. Telephone: 214469627. Fax: 214411277. w-mail: [email protected]
Rights & Permissions [Opens in a new window]

Summary

To identify transposons that may be of use for mutagenesis we investigated the genetic molecular basis of a case of flower colour variegation in Linaria, a close relative of the model species Antirrhinum majus. We show that this variegation is attributable to an unstable mutant allele of the gene encoding dihydroflavonol-4-reductase, one of the enzymes required for anthocyanin biosynthesis. This allele carries an insertion of a transposon belonging to the CACTA family (Tl1, Transposon Linaria 1) which blocks its expression thus conferring an ivory flower colour phenotype. Tl1 is occasionally excised in dividing epidermal cells to produce clonal patches of red tissue on the ivory background, and in cells giving rise to gametes to generate reversion alleles conferring a fully coloured phenotype. This finding may open the way for targeted transposon-mutagenesis in Linaria, and hence for using this genus in comparative genetic studies.

Type
Research Article
Information
Genetics Research , Volume 89 , Issue 3 , June 2007 , pp. 129 - 134
Copyright
Copyright © Cambridge University Press 2007

1. Introduction

Although transposons have been identified in numerous species as DNA segments with particular structures, in only a few cases has it been possible to detect phenotypic consequences of their activity. Such consequences are particularly evident in the case of plant transposons which, when inserted in pigmentation genes, give rise to unstable mutant alleles conferring variegated colour phenotypes (Bonas et al., Reference Bonas, Sommer and Saedler1984; Fedoroff et al., Reference Fedoroff, Furtek and Nelson1984; Kroon et al., Reference Kroon, Souer, de Graaff, Xue, Mol and Koes1994; Inagaki et al., Reference Inagaki, Hisatomi, Suzuki, Kasahara and Iida1994; Chopra et al., Reference Chopra, Brendel, Zhang, Axtell and Peterson1999). In maize, Antirrhinum and Petunia, detailed analysis of alleles of this type has led to the use of transposons as tools for systematically generating mutants (Carpenter & Coen, Reference Carpenter and Coen1990; Luo et al., Reference Luo, Coen, Doyle and Carpenter1991; van Houwelingen et al., Reference May, Liu, Vollbrecht, Senior, Rabinowicz, Roh, Pan, Stein, Freeling, Alexander and Martienssen1998; May & Martienssen, Reference May and Martienssen2003). An advantage of transposon mutagenesis is that it offers the prospect of isolating genes from unstable alleles or, conversely, selecting for mutants carrying transposon insertions or transposon-induced deletions in known genes (Koes et al., Reference Koes, Souer, van Houwelingen, Mur, Spelt, Quatrocchio, Wing, Oppedijk, Ahmed, Maes, Gerats, Hoogeveen, Meesters, Kloos and Mol1995; Ingram et al., Reference Ingram, Doyle, Carpenter, Schultz, Simon and Coen1997; Davies et al., Reference Davies, Motte, Keck, Saedler, Sommer and Schwarz-Sommer1999; Galego & Almeida, Reference Galego and Almeida2002; May et al., Reference May, Liu, Vollbrecht, Senior, Rabinowicz, Roh, Pan, Stein, Freeling, Alexander and Martienssen2003). Furthermore, transposition can in some instances be exploited to genetically mark clones of cells, hence for lineage analysis or for studying cell autonomy in gene action (Dawe & Freeling, Reference Dawe and Freeling1990; Dudley & Poethig, Reference Dudley and Poethig1993; Carpenter & Coen, Reference Carpenter and Coen1995; Hantke et al., Reference Hantke, Carpenter and Coen1995; Vincent et al., Reference Vincent, Carpenter and Coen1995; Rolland-Lagan et al., Reference Rolland-Lagan, Bangham and Coen2003).

The possibilities above have been exploited for detailed studies on the evolution of genetic and developmental mechanisms incorporating comparisons of mutants in diverse, distantly related, model species. For example, site-directed selection for transposon insertions has been used to investigate the role of Antirrhinum counterparts of the Arabidopsis Ap2 gene in flower development (Keck et al., Reference Keck, McSteen, Carpenter and Coen2003). Equivalent tools are, however, unavailable for comparisons between Antirrhinum and its close relatives. Thus, as an exploratory step towards expanding transposon mutagenesis to a genus closely related to Antirrhinum, we have investigated the genetic molecular basis of a case of flower colour variegation in Linaria, a member of the Antirrhinum tribe (Olmstead et al., Reference Olmstead, dePamphilis, Wolfe, Young, Elisens and Reeves2001; Vargas et al., Reference Vargas, Rossello, Oyama and Güemes2004). We show that such variegation results from activity of a CACTA transposon inserted in the gene encoding dihydroflavonol-4-reductase and that Tl1 transposes in cells giving rise to gametes.

2. Materials and methods

(i) Origin of plants

The Linaria plants considered here (Fig. 1) were found in a garden population of unknown origin. Many morphological and colour variants occurred in the population, which could possibly comprise more than one species or species hybrids. Assigning individuals to particular species was therefore problematic. One possibility was that plants with variegated colour belonged to or comprised part of Linaria maroccanna, a species from which popular garden varieties are derived. These plants had reniform seed with distinct transverse ridges and a style with lateral lobes bearing discrete stigmatic areas, all traits common to L. maroccanna and a few other species (Sutton, Reference Sutton1988). However, it was unclear whether capsules had conspicuously unequal loculi, a trait which, together with those above, is common to only L. maroccana and L. gharbensis.

Fig. 1. Flower colour phenotypes in Linaria. A fully red inflorescence is shown on the left and variegated inflorescence on the right.

(ii) Primers

The following primers were used:

  • i1: AA(A/G)(C/A)GNCA(C/T)ACNGA(C/T)CCNGGNACNAT(A/C/T)AC

  • i2: TTNACNGC(T/C)TG(G/A)TG(G/A)TCNGC(G/A)TT(T/C)TT

  • p1: GGNTT(C/T)AT(A/C/T)GGN(A/T)(G/C)NTGG(T/C)TNGTNATG

  • p2: AA(G/A)ATGACNGGNTGGATGTA(C/T)TT

  • p3: GGTCACTGGAATCGGTTTCATCG

  • p4: GTGGCTATACCATTCGTGCAAC

  • p5: CTCATGACCAGCCATGAGCCAA

  • p6: CTGCCAACGTGACGAACTTCAC

  • p7: CCACAATAACATGTCCGTATC

  • p8: TGCAACTCACAATTAGCT

  • p9: CCGTTGGCTCCAGTAACGCAT

  • p10: GCGATACTAGGCAACAAGTTC

  • Tam1-3′: CCAGAGCCCAATTTTGTTGTAGTG

The positions and orientations of primers p1 to p9 are indicated with arrowheads in Fig. 2C.

Fig. 2. Molecular analysis of colour variegation. (A) Northern blot of RNA from inflorescences of a plant with variegated color (var) and a fully red revertant (red) probed with LPAL and LINC. The two plants were sibs derived from a cross between variegated plants. The same result was obtained in all of four independent comparisons with pairs of variegated and revertant sibs. (B) Southern blot of DNA from genotypes indicated above the lanes. DNAs were cut with HindIII and probed with a full-length cDNA segment of LPAL. Sizes of fragments in kilobases are indicated on the left. Lpal-1/LPAL was derived from a cross between two homozygotes for Lpal-1. The LPAL/LPAL DNA was from a red segregant in a family derived from crossing two independent revertants. (C) Structure of the LPAL gene. Transcription is from left to right with UTRs in dark grey, coding region in white and introns in light grey. The triangle represents Tl1 and the arrowheads indicate primers used in this work. H indicates sites cut by HindIII. (D) Sequences of Lpal alleles in the vicinity of the Tl1 insertion site. The TSD is in bold and Tl1 ends in italic. Reversion alleles (Revs.) 2, 3 and 4 are germinal events. Sequence 5 is from a large somatic sector. Sequence 1 was found in four independent germinal events and in two large somatic sectors.

(iii) Molecular analysis

cDNA Segments of LPAL and LINC were isolated by RT-PCR using RNA extracted from red inflorescences and degenerate primers designed according to aligned sequences of genes encoding dihydroflavonol-4-reductase (p1 and p2; 1 and 2 in Fig. 2C) and flavonol-3-hydroxylase (i1 and i2) from various species. A genomic segment containing the right end of Tl1 and its flanking Lpal sequence was isolated by PCR amplification of DNA from Lpal-1 homozygotes using an LPAL-specific primer, p3, and a primer for the 3′ end of the Antirrhinum transposon Tam1 (Tam1–3′) at low annealing temperature (55°C). A genomic segment comprising 478 bp of the right end of Tl1 and its flanking Lpal-2 region was then obtained by inverse PCR using p4 and p5 on DNA cut with Sau3A prior to ligation. Using the same approach on DNA from fully red revertants resulted in amplification of a segment spanning the site of Tl1 excision and a further 453 bp to its left. A segment containing 342 bp of the left end of Tl1 was then obtained by inverse PCR using p6 and p7 on DNA from Lpal-1 homozygotes cut with Sau3A prior to ligation. The LPAL transcription start site was determined by 5′RACE using p3 and p5. A nearly full length cDNA was then obtained by 3′-RACE using p8 as specific primer. A genomic segment spanning the transcribed region was amplified using p8 and p10. Segments described above were cloned in pBluescript SK (Stratagene) and sequenced automatically (Stabvida, Oeiras). Segments from reversion alleles, carrying Tl1 excision footprints, were obtained by PCR using p5 and p6. Sequences of these segments were then determined directly on PCR products using p9.

Sequences for LPAL, Tl1 and LINC have been deposited in the GenBank Data Library under accession nos. EF517133, EF517134 and EF517135, respectively.

3. Results and discussion

(i) Germinal instability of flower colour

Plants with variegated colour had ivory flowers and green stems with clonal patches of red and purple cells respectively, a phenotype very similar to those conferred by some unstable alleles of pigmentation genes in Antirrhinum (Coen & Carpenter, Reference Coen and Carpenter1986; Schwarz-Sommer et al., Reference Schwarz-Sommer, Davies and Hudson2003; Fig. 1). Typically, insertion of a transposon in a pigmentation gene blocks gene expression giving non-pigmented tissue. However, the transposon is occasionally excised from the gene in dividing cells, hence giving clones of pigmented cells on the non-pigmented background. In addition, the transposon can excise in cells giving rise to gametes and therefore a few plants with wild-type phenotype (revertants) can be recovered in progenies of homozygotes for the unstable allele. To test whether the apparent instability in Linaria occurred in germinal tissue we examined pigmentation phenotypes in progenies from crosses between plants with variegated colour (these plants were self-incompatible). In nearly 700 progeny from five crosses, 10 plants had fully red flowers and purple stems whereas the remaining ones had the parental phenotype. Thus, by analogy to Antirrhinum, an explanation for the variegated colour phenotype and genetic behaviour of Linaria plants was that they carried an active transposon in a pigmentation gene.

(ii) Lpal-1: an unstable mutant allele of the gene encoding dihydroflavonol-4-reductase

In Antirrhinum, red colour depends on structural genes encoding enzymes required for anthocyanin synthesis, and on regulatory genes that control the expression of structural genes (Almeida et al., Reference Almeida, Carpenter, Robbins, Martin and Coen1989; Goodrich et al., Reference Goodrich, Carpenter and Coen1992; Schwinn et al., Reference Schwinn, Venail, Shang, Mackay, Alm, Butelli, Oyama, Bailey, Davies and Martin2006). Null mutants for structural genes are in general devoid of anthocyanin pigment in all plant parts whereas regulatory mutants show altered colour patterns relative to wild-type. In Linaria, except for the clonal patches of pigmented cells, all plant parts lacked red pigment, suggestive of mutation in a structural gene. Null mutations affecting enzymes required at early steps of the anthocyanin synthesis pathway in Antirrhinum, such as chalcone synthase, result in albino flowers whereas mutations affecting late steps give ivory flower colour identical to that in Linaria (see Coen et al., Reference Coen, Robbins, Almeida, Hudson, Carpenter, Berg and Howe1989). Late steps include those requiring flavonol-3-hydroxylase and dihydroflavonol-4-reductase, the products of the INCOLORATA (INC) and PALLIDA (PAL) genes respectively (Martin et al., Reference Martin, Carpenter, Sommer, Saedler and Coen1985, Reference Martin, Prescott, Mackay, Bartlett and Vrijlandt1991; Coen et al., Reference Coen, Carpenter and Martin1986).

To determine whether variegated plants were mutant at the counterparts of either INC or PAL (LINC or LPAL) we probed Northern blots of inflorescence RNA from variegated and fully red sibs derived from the crosses described above with cDNA segments for the LINC or LPAL genes (Fig. 2A). LINC mRNA was detected in both variegated and red inflorescences whereas LPAL mRNA was detected in red but not in variegated inflorescences, suggesting that mutation at Lpal caused colour variegation. Consistent with this and with Lpal carrying a transposon insertion that could excise in germinal tissue, Southern analysis showed that progeny with variegated colour derived from the crosses above were homozygous for an Lpal allele (Lpal-1) whereas all their fully red sibs tested (six from the five families) were heterozygous for Lpal-1 and putative reversion alleles (Fig. 2B).

(iii) Lpal-1 carries a CACTA transposon (Tl1, Transposon Linaria 1) in the 5′-UTR

Transposons of two classes, distinguishable according to the sequences of their terminal inverted repeats (TIR) and sizes of their target site duplications (TSD), give rise to unstable alleles in Antirrhinum. One class includes Tam3 (the same family as Ac/Ds in maize; Hehl et al., Reference Hehl, Nacken, Krause, Saedler and Sommer1991) and the other class includes a number of transposons, such as Tam1, which together with other plant transposons (e.g. Spm in maize) belong to a family known as CACTA, after a common sequence motif at their TIRs (Nacken et al., Reference Nacken, Piotowiak, Saedler and Sommer1991; Kunze & Weil, Reference Kunze, Weil and Craig2002). To determine whether Lpal-1 carries a transposon of one of these classes we amplified DNA from Lpal-1 homozygotes using primers for LPAL and for the ends of either Tam3 or Tam1. This resulted in amplification of DNA with a Tam1 primer. Full characterization of the Lpal-1 allele then showed that it carries a CACTA transposon (Tl1, Transposon Linaria 1) in the 5′UTR, 44 bp upstream of the translation initiation codon and 29 bp downstream of the transcription start site (Fig. 2C). Similarity searches showed that the 342 bp sequence of the left end of Tl1 has over 60% identity with Tam1, Tam2 and Tam4, all Antirrhinum CACTA transposons.

To test further whether Tl1 was responsible for Lpal-1 instability, we took advantage of the property of imprecise transposon excision (Saedler & Nevers, Reference Saedler and Nevers1985; Coen et al., Reference Coen, Carpenter and Martin1986, Reference Coen, Robbins, Almeida, Hudson, Carpenter, Berg and Howe1989). Duplications of short sequences at host loci generated on integration (TSD) can be retained intact or in diverse modified forms following excision. In addition, transposon excision can generate deletions adjacent to the TSD. As is typical for CACTA transposons, Tl1 was flanked by a 3 bp TSD in Lpal-1 (the trinucleotide TAA, which had a single copy in plants with red flowers of unknown relatedness to the variegated plants; Fig. 2D). In germinal reversion alleles and in single large somatic red clones from which DNA could be extracted we found no cases with a single TAA sequence, six independent cases in which both copies of the TSD were retained intact, three cases with modified TSDs and one case in which a 10 bp deletion was generated adjacent to the TSD (Fig. 2D). Presumably, these small sequence modifications did not have a large effect on LPAL expression given their location in the 5′UTR.

(iv) Concluding remarks

In conclusion, by applying knowledge derived from the analysis of unstable pigmentation mutants in Antirrhinum to a case of flower colour variegation in Linaria we have trapped an active copy of a transposon belonging to the CACTA family in the gene encoding dihydroflavonol-4-reductase. This may open the way for developing a system to select for transposon insertions in Linaria counterparts of Antirrhinum genes such as, for example, those controlling flower morphology. The two genera have readily comparable flowers with similar structures and overall dorsoventral patterns (Cubas et al., Reference Cubas, Vincent and Coen1999; Gübitz et al., Reference Gübitz, Caldwell and Hudson2003). However, they differ in details of petal size and shape as well as in some conspicuous traits. The most obvious is a long spur in the ventral petal of Linaria which is thought to be the natural counterpart of ventral petal outgrowths arising in some gain-of-function Antirrhinum mutants (Golz et al., Reference Golz, Keck and Hudson2002). Thus, Antirrhinum and Linaria might provide a useful pair of close relatives for investigating origins of both subtle and large morphological variations.

References

Almeida, J., Carpenter, R., Robbins, T. P., Martin, C. & Coen, E. S. (1989). Genetic interactions underlying flower color patterns in Antirrhinum majus. Genes and Development 3, 1758–1767.CrossRefGoogle ScholarPubMed
Bonas, U., Sommer, H. & Saedler, H. (1984). The 17 Kb Tam1 element of Antirrhinum majus induces a 3 bp duplication upon integration into the chalcone synthase gene. EMBO Journal 3, 1015–1019.CrossRefGoogle ScholarPubMed
Carpenter, R. & Coen, E. S. (1990). Floral homeotic mutations produced by transposon mutagenesis in Antirrhinum majus. Genes and Development 4, 1483–1493.CrossRefGoogle ScholarPubMed
Carpenter, R. & Coen, E. S. (1995). Transposon induced chimeras show that FLORICAULA, a meristem identity gene, acts non-autonomously between cell-layers. Development 121, 19–26.CrossRefGoogle ScholarPubMed
Chopra, S., Brendel, V., Zhang, J. B., Axtell, J. D. & Peterson, T. (1999). Molecular characterization of a mutable pigmentation phenotype and isolation of the first active transposable element from Sorghum bicolor. Proceedings of the National Academy of Sciences of the USA 96, 15330–15335.CrossRefGoogle ScholarPubMed
Coen, E. S. & Carpenter, R. (1986). Transposable elements in Antirrhinum majus: generators of genetic diversity. Trends in Genetics 2, 292–296.CrossRefGoogle Scholar
Coen, E. S., Carpenter, R. & Martin, C. (1986). Transposable elements generate novel spatial patterns of gene expression in Antirrhinum. Cell 47, 285–296.CrossRefGoogle ScholarPubMed
Coen, E. S., Robbins, T. P., Almeida, J., Hudson, A. & Carpenter, R. (1989). Consequences and mechanisms of transposition in Antirrhinum majus. In Mobile DNA (ed. Berg, D. E. & Howe, M. M.), pp. 413–436. Washington, DC: American Society for Microbiology.Google Scholar
Cubas, P., Vincent, C. & Coen, E. S. (1999). An epigenetic mutation responsible for natural variation in floral symmetry. Nature 401, 157–161.CrossRefGoogle ScholarPubMed
Davies, B., Motte, P., Keck, E., Saedler, H., Sommer, H. & Schwarz-Sommer, Zs. (1999). PLENA and FARINELLI: redundancy and regulatory interactions between two Antirrhinum MADS-box factors controlling flower development. EMBO Journal 18, 4023–4034.CrossRefGoogle ScholarPubMed
Dawe, R. K. & Freeling, M. (1990). Clonal analysis of the cell lineages in the male flower of maize. Developmental Biology 142, 233–245.CrossRefGoogle ScholarPubMed
Dudley, M. & Poethig, R. S. (1993). The heterochronic Teopod1 and Teopod2 mutations of maize are expressed non-cell-autonomously. Genetics 133, 389–399.CrossRefGoogle ScholarPubMed
Fedoroff, N., Furtek, D. & Nelson, O. (1984). Cloning of the Bronze locus in maize by a simple and generalizable procedure using the transposable controlling element Ac. Proceedings of the National Academy of Sciences of the USA 81, 3825–3829.CrossRefGoogle Scholar
Galego, L. & Almeida, J. (2002). Role of DIVARICATA in the control of dorsoventral asymmetry in Antirrhinum flowers. Genes and Development 16, 880–891.CrossRefGoogle ScholarPubMed
Golz, J. F., Keck, E. J. & Hudson, A. (2002). Spontaneous mutations in KNOX genes give rise to a novel flower structure in Antirrhinum. Current Biology 12, 515–522.CrossRefGoogle ScholarPubMed
Goodrich, J., Carpenter, R. & Coen, E. S. (1992). A common gene regulates pigmentation pattern in diverse plant species. Cell 6, 955–964.CrossRefGoogle Scholar
Gübitz, T., Caldwell, A. & Hudson, A. (2003). Rapid molecular evolution of CYCLOIDEA-like genes in Antirrhinum and its relatives. Molecular Biology and Evolution 20, 1537–1544.CrossRefGoogle ScholarPubMed
Hantke, S., Carpenter, R. & Coen, E. S. (1995). Expression of FLORICAULA in single-cell layers of periclinal chimeras activates downstream genes in all layers of floral meristems. Development 121, 27–35.CrossRefGoogle ScholarPubMed
Hehl, R., Nacken, W. K., Krause, A., Saedler, H. & Sommer, H. (1991). Structural analysis of Tam3, a transposon from Antirrhinum majus, reveals homologies to the Ac element of maize. Plant Molecular Biology 16, 369–371.CrossRefGoogle Scholar
Inagaki, Y., Hisatomi, Y., Suzuki, T., Kasahara, K. & Iida, S. (1994). Isolation of a Suppressor-mutator Enhancer-like transposable element, TPN1, from Japanese Morning Glory bearing variegated flowers. Plant Cell 6, 375–383.Google ScholarPubMed
Ingram, G., Doyle, S., Carpenter, R., Schultz, E., Simon, R. & Coen, E. S. (1997). Dual role for fimbriata in regulating floral homeotic genes and cell division in Antirrhinum. EMBO Journal 16, 6521–6534.CrossRefGoogle ScholarPubMed
Keck, E., McSteen, P., Carpenter, R. & Coen, E. S. (2003). Separation of genetic functions controlling organ identity in flowers. EMBO Journal 22, 1058–1066.CrossRefGoogle ScholarPubMed
Koes, R., Souer, E., van Houwelingen, A., Mur, L., Spelt, C., Quatrocchio, F., Wing, J., Oppedijk, B., Ahmed, S., Maes, T., Gerats, T., Hoogeveen, P., Meesters, M., Kloos, D. & Mol, J. N. M. (1995). Targeted gene inactivation in Petunia by PCR-based selection of transposon insertion mutants. Proceedings of the National Academy of Sciences of the USA 92: 8149–8153.CrossRefGoogle ScholarPubMed
Kroon, J., Souer, E., de Graaff, A.Xue, Y., Mol, J. & Koes, R. (1994). Cloning and structural analysis of the anthocyanin pigmentation locus Rt of Petunia hybrida: characterization of the insertion sequences in two mutant alleles. Plant Journal 5: 69–80.CrossRefGoogle ScholarPubMed
Kunze, R. & Weil, C. F. (2002). The hAT and CACTA superfamily of plant transposons. In Mobile DNA (ed. Craig, N. L.), pp. 565–610. Washington, DC: American Society for Microbiology.Google Scholar
Luo, D., Coen, E. S., Doyle, S. & Carpenter, R. (1991). Pigmentation mutants produced by transposon mutagenesis in Antirrhinum majus. Plant Journal 1, 59–69.CrossRefGoogle ScholarPubMed
Martin, C., Carpenter, R., Sommer, H., Saedler, H. & Coen, E. S. (1985). Molecular analysis of instability in flower pigmentation in Antirrhinum majus following isolation of the pallida locus by transposon tagging. EMBO Journal 4, 1625–1630.CrossRefGoogle ScholarPubMed
Martin, C., Prescott, A., Mackay, S., Bartlett, J. & Vrijlandt, E. (1991). Control of anthocyanin biosynthesis in flowers of Antirrhinum majus. Plant Journal 1, 37–49.CrossRefGoogle ScholarPubMed
May, B. & Martienssen, R. (2003). Transposon mutagenesis in the study of plant development. Critical Reviews in Plant Sciences 22, 1–35.CrossRefGoogle Scholar
May, B. P., Liu, H., Vollbrecht, E., Senior, L., Rabinowicz, P. D., Roh, D., Pan, X. K., Stein, L., Freeling, M.Alexander, D. & Martienssen, R. (2003). Maize-targeted mutagenesis: a knockout resource for maize. Proceedings of the National Academy of Sciences of the USA 100, 11541–11546.CrossRefGoogle ScholarPubMed
Nacken, W., Piotowiak, R., Saedler, H. & Sommer, H. (1991). The transposable element Tam1 from Antirrhinum majus shows structural homology to the maize transposon En/Spm and has no sequence specificity of insertion. Molecular General Genetics 228, 201–208.CrossRefGoogle Scholar
Olmstead, R. G., dePamphilis, C., Wolfe, A. D., Young, N. D., Elisens, W. J. & Reeves, P. A. (2001). Disintegration of Scrophulariaceae. American Journal of Botany 88, 348–361.CrossRefGoogle ScholarPubMed
Rolland-Lagan, A., Bangham, J. & Coen, E. S. (2003). Growth dynamics underlying petal shape and asymmetry. Nature 422, 161–163.CrossRefGoogle ScholarPubMed
Saedler, H. & Nevers, P. (1985). Transposition in plants: a molecular model. EMBO Journal 4, 585–590.CrossRefGoogle ScholarPubMed
Schwarz-Sommer, Z., Davies, B. & Hudson, A. (2003). An everlasting pioneer: the story of Antirrhinum research. Nature Reviews Genetics 4, 657–666.CrossRefGoogle ScholarPubMed
Schwinn, K., Venail, J., Shang, Y., Mackay, S., Alm, V., Butelli, E., Oyama, R., Bailey, P., Davies, K. & Martin, C. (2006). A small family of MYB-regulatory genes controls floral pigmentation intensity and patterning in the genus Antirrhinum. Plant Cell 18, 831–851.CrossRefGoogle ScholarPubMed
Sutton, D. A. (1988). A revision of the tribe Antirrhineae. London: Oxford University Press.Google Scholar
Van Houwelingen, A., Souer, E., Spelt, K., Kloos, D., Mol, J. & Koes, R. (1998). Analysis of flower pigmentation mutants generated by random transposon mutagenesis in Petunia hybrida. Plant Journal 13, 39–50.Google ScholarPubMed
Vargas, P., Rossello, J. A., Oyama, R. & Güemes, J. (2004). Molecular evidence for naturalness of genera in the tribe Antirrhineae (Scrophulariaceae) and three independent evolutionary lineages in the New World and the Old. Plant Systematics and Evolution 249, 151–172.CrossRefGoogle Scholar
Vincent, C., Carpenter, R. & Coen, E. S. (1995). Cell lineage patterns and homeotic gene activity during Antirrhinum flower development. Current Biology 5, 1449–1458.CrossRefGoogle ScholarPubMed
Figure 0

Fig. 1. Flower colour phenotypes in Linaria. A fully red inflorescence is shown on the left and variegated inflorescence on the right.

Figure 1

Fig. 2. Molecular analysis of colour variegation. (A) Northern blot of RNA from inflorescences of a plant with variegated color (var) and a fully red revertant (red) probed with LPAL and LINC. The two plants were sibs derived from a cross between variegated plants. The same result was obtained in all of four independent comparisons with pairs of variegated and revertant sibs. (B) Southern blot of DNA from genotypes indicated above the lanes. DNAs were cut with HindIII and probed with a full-length cDNA segment of LPAL. Sizes of fragments in kilobases are indicated on the left. Lpal-1/LPAL was derived from a cross between two homozygotes for Lpal-1. The LPAL/LPAL DNA was from a red segregant in a family derived from crossing two independent revertants. (C) Structure of the LPAL gene. Transcription is from left to right with UTRs in dark grey, coding region in white and introns in light grey. The triangle represents Tl1 and the arrowheads indicate primers used in this work. H indicates sites cut by HindIII. (D) Sequences of Lpal alleles in the vicinity of the Tl1 insertion site. The TSD is in bold and Tl1 ends in italic. Reversion alleles (Revs.) 2, 3 and 4 are germinal events. Sequence 5 is from a large somatic sector. Sequence 1 was found in four independent germinal events and in two large somatic sectors.